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Characterization of a lytic polysaccharide monooxygenase from Aspergillus fumigatus shows functional variation among family AA11 fungal LPMOs

Open AccessPublished:November 16, 2021DOI:https://doi.org/10.1016/j.jbc.2021.101421
      The discovery of oxidative cleavage of recalcitrant polysaccharides by lytic polysaccharide monooxygenases (LPMOs) has affected the study and industrial application of enzymatic biomass processing. Despite being widespread in fungi, LPMOs belonging to the auxiliary activity (AA) family AA11 have been understudied. While these LPMOs are considered chitin active, some family members have little or no activity toward chitin, and the only available crystal structure of an AA11 LPMO lacks features found in bacterial chitin-active AA10 LPMOs. Here, we report structural and functional characteristics of a single-domain AA11 LPMO from Aspergillus fumigatus, AfAA11A. The crystal structure shows a substrate-binding surface with features resembling those of known chitin-active LPMOs. Indeed, despite the absence of a carbohydrate-binding module, AfAA11A has considerable affinity for α-chitin and, more so, β-chitin. AfAA11A is active toward both these chitin allomorphs and enhances chitin degradation by an endoacting chitinase, in particular for α-chitin. The catalytic activity of AfAA11A on chitin increases when supplying reactions with hydrogen peroxide, showing that, like LPMOs from other families, AfAA11A has peroxygenase activity. These results show that, in stark contrast to the previously characterized AfAA11B from the same organism, AfAA11A likely plays a role in fungal chitin turnover. Thus, members of the hitherto rather enigmatic family of AA11 LPMOs show considerable structural and functional differences and may have multiple roles in fungal physiology.

      Keywords

      Abbreviations:

      AA (auxiliary activity), AscA (ascorbic acid), H2O2 (hydrogen peroxide), LPMO (lytic polysaccharide monooxygenase), MWCO (molecular weight cutoff), O2 (oxygen), PASC (phosphoric acid-swollen cellulose), PES (polyether sulfone), ChiC (SmChiC)
      Many microorganisms produce an arsenal of carbohydrate-active enzymes that concertedly operate to degrade carbohydrate biomass. These enzyme systems are fundamental to the global carbon cycle and prevent an environmental buildup of stable and abundant sugar polymers such as cellulose and chitin (
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      ). In the case of an O2-driven reaction, the reaction requires two externally delivered electrons per catalytic cycle (
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      ). Cellulose-active LPMOs oxidize either the C1 or the C4 carbon of the scissile glycosidic bond; some LPMOs exclusively act on C1 or C4, whereas others are less specific and produce a mixture of C1-oxidized and C4-oxidized products (
      • Beeson W.T.
      • Phillips C.M.
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      ). This oxidation leads to spontaneous subsequent bond cleavage (
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      Cellobiose dehydrogenase and a copper-dependent polysaccharide monooxygenase potentiate cellulose degradation by Neurospora crassa.
      ). C6-oxidized products have also been reported (
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      ), but their importance and connection to the chain-cleaving function of LPMOs remain unclear. For chitin-active LPMOs, only C1-oxidizing activity has been reported. The latter activity leads to the generation of new chain ends that are lactones, which spontaneously convert to aldonic acids (
      • Vaaje-Kolstad G.
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      • Liu Z.
      • Zhai H.
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      ).
      Despite a strictly conserved catalytic center, that is, the histidine brace, LPMOs display large sequence diversity and currently populate eight “auxiliary activity” (AA) families in the Carbohydrate-Active enZymes database (http://www.cazy.org/Auxiliary-Activities.html; as of August 2021), a sequence-based database of carbohydrate-active enzymes, namely AA families 9 to 11 and 13 to 17 (
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      • Lombard V.
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      ). Fungal LPMOs appear in families 9, 11, 13, 14, and 16. Of these, the cellulose-active fungal AA9 LPMOs are the most abundant and the best studied. Remarkably, while chitin-active LPMOs are commonly found among members of the AA10 family, which lacks fungal enzymes, and while fungi are known for their ability to produce multiple chitinases and degrade chitin (
      • Seidl V.
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      A complete survey of Trichoderma chitinases reveals three distinct subgroups of family 18 chitinases.
      ), little is known about fungal chitin-active LPMOs.
      Fungal chitin-active LPMOs have been classified into the AA11 family (
      • Hemsworth G.R.
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      • Davies G.J.
      • Walton P.H.
      Discovery and characterization of a new family of lytic polysaccharide monooxygenases.
      ). So far, only three members of this family have been subjected to some degree of functional characterization (
      • Hemsworth G.R.
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      ). Available structural information is limited to the incomplete crystal structure of the catalytic domain of a two-domain AA11 from Aspergillus oryzae (AoAA11; (
      • Hemsworth G.R.
      • Henrissat B.
      • Davies G.J.
      • Walton P.H.
      Discovery and characterization of a new family of lytic polysaccharide monooxygenases.
      )). Family 11 LPMOs are the most widespread of fungal LPMOs as they occur in Dikarya and most basal fungal lineages (
      • Várnai A.
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      Fungal lytic polysaccharide monooxygenases (LPMOs): Biological importance and applications.
      ). The ectomycorrhizal fungus Choiromyces venosus contains the highest number of genes (i.e., 14 genes) encoding AA11 LPMOs. AA11s are most prevalent in Ascomycota, where at least one gene encoding an AA11 LPMO has been found in 385 (99.2%) out of the 388 analyzed genomes, and where the average number of genes encoding AA11s is 3.9 ± 1.8 (
      • Várnai A.
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      • Horn S.J.
      • Eijsink V.G.H.
      • Berrin J.-G.
      Fungal lytic polysaccharide monooxygenases (LPMOs): Biological importance and applications.
      ). The genome of the ascomycete Aspergillus fumigatus (
      • Nierman W.C.
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      • Berriman M.
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      Genomic sequence of the pathogenic and allergenic filamentous fungus Aspergillus fumigatus.
      ) contains three genes encoding family AA11 LPMOs, here referred to as AfAA11A (UniProt ID: Q4WF00; gene, AFUA_3G03950), a single domain enzyme, AfAA11B (UniProt ID: Q4WEH3; gene, AFUA_5G03010), a two-domain enzyme with a C-terminal domain of unknown function, and AfAA11C (UniProt ID: Q4WM72; gene, AFUA_6G10930), also a two-domain enzyme with a C-terminal domain of unknown function. Interestingly, a recent in-depth functional study of AfAA11B (
      • Rieder L.
      • Petrović D.
      • Väljamäe P.
      • Eijsink V.G.H.
      • Sørlie M.
      Kinetic characterization of a putatively chitin-active LPMO reveals a preference for soluble substrates and absence of monooxygenase activity.
      ), which is very similar to AoAA11 (73% sequence identity for the catalytic domains), showed that this enzyme has low activity on chitin (i.e., much lower than chitin-active bacterial AA10 LPMOs) while being efficient when acting on soluble substrates. This raises questions as to the natural function of AA11 LPMOs.
      To gain more insight into the AA11 LPMO family and oxidative chitin conversion by fungi, we have structurally and functionally characterized AfAA11A, including studies of this protein's ability to potentiate the action of chitinases. We also assessed whether this AA11 LPMO, like AA9 and AA10 LPMOs, is capable of employing H2O2 as its oxygen-containing cosubstrate. Next to providing insight into the functionality of family AA11 LPMOs, our study provides the first complete crystal structure of a member of this family.

      Results

      Protein production and deglycosylation

      The gene encoding AfAA11A (UniProt ID: Q4WF00) was cloned in Pichia pastoris, and recombinant AfAA11A was purified to homogeneity from the culture supernatant using hydrophobic interaction chromatography and size-exclusion chromatography (Fig. 1). SDS-PAGE analysis showed that purified AfAA11A had a mass close to 30 kDa, indicating that the protein carried glycan moieties. Analysis with the NetNGlyc server (http://www.cbs.dtu.dk/services/NetNGlyc/) showed one possible N-glycosylation site, at Asn62. Indeed, treatment of the purified protein with an endo-β-N-acetylglucosaminidase from Enterococcus faecalis (EfEndo18A) (
      • Bohle L.A.
      • Mathiesen G.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      An endo-ss-N-acetylglucosaminidase from Enterococcus faecalis V583 responsible for the hydrolysis of high-mannose and hybrid-type N-linked glycans.
      ) led to a reduction in the protein mass to a value, approximately 22 kDa, that corresponds well with the theoretical mass (21,750 Da; Fig. 1). Initial comparison of the glycosylated and deglycosylated protein batches indicated similar activities and properties, although minor, borderline significant, differences were observed in, for example, substrate binding and melting point (not shown). Here, we report the functional characterization of the glycosylated protein.
      Figure thumbnail gr1
      Figure 1SDS-PAGE analysis of purified AfAA11A with or without deglycosylation. SDS-PAGE was done with stain-free precast gels and fluorescence imaging. In both panels, lane 1 contains the BenchMark protein ladder. In panel A, lanes 2 and 3 show two batches of the N-glycosylated protein, whereas lane 4 shows the protein after overnight deglycosylation with EfEndo18A and subsequent removal of EfEndo18A by chromatography. Panel B shows how deglycosylation proceeded in the initial phase of the reaction with EfEndo18A; lane 2, untreated AfAA11A; lanes 3 to 5, the protein after treatment with EfEndo18A for 5, 10, and 15 min, respectively. Note that glycosylated AfAA11A and EfEndo18A have similar masses and are not separated on this gel.

      AfAA11A crystal structure

      Crystallization conditions were screened for both N-glycosylated and de-N-glycosylated AfAA11A. Only one of the conditions tested resulted in a protein crystal suitable for structure analysis, for the de-N-glycosylated protein. The structure of AfAA11A was solved by X-ray crystallography, and a complete structure was obtained, which, however, lacked a bound metal ion in the histidine brace. Crystal data, as well as data collection and refinement statistics, are provided in Table 1. Of note, except for one GlcNAc attached to Asn62, the structure of this N-deglycosylated protein showed no other signs of N-glycosylation or O-glycosylation. Asn62 is not part of the flat LPMO surface containing the catalytic site and is located far away from the catalytic histidines; the Cβ–Cβ distances for Asn62 and His1 and His71 are 21.3 and 22.2 Å, respectively, with side chains pointing into opposite directions.
      Table 1Crystal data, data collection, and refinement statistics
      Crystal data
      AfAA11A
       Space groupP 212121
       Crystal parametersa = 40.99 Å, b = 47.28 Å, and c = 115.59 Å
      α = 90°, β = 90°, and γ = 90°
      Data collection
       X-ray sourceMAXIV, BioMAX
       Resolution (Å)
      Values for outer shell are given in parenthesis.
      29.89–1.50 (1.53–1.50)
       Wavelength (Å)0.946444
       Temperature (K)100
       Number of unique reflections36,807 (1849)
       Completeness
      Values for outer shell are given in parenthesis.
      99.8 (99.8)
       Redundancy
      Values for outer shell are given in parenthesis.
      12.3 (8.3)
       CC half
      Values for outer shell are given in parenthesis.
      0.999 (0.986)
      I/σ(I)
      Values for outer shell are given in parenthesis.
      19.2 (3.5)
      Rmerge
      Rmerge=∑|I−⟨I⟩|∑I.
      0.061 (0.361)
      Refinement statistics
      Rcryst
      Rcryst=∑(|Fobs|−|Fcalc|)∑|Fobs|.
      0.146
      Rfree
      Rfree is the Rcryst value calculated on the 5% reflections excluded for refinement.
      0.197
       Wilson B-factor (Å2)28.9
       Ramachandran plot, in most favored/other allowed regions (%)98/2
       Added waters182
       Protein Data Bank code7P3U
      a Values for outer shell are given in parenthesis.
      b Rmerge=|II|I.
      c Rcryst=(|Fobs||Fcalc|)|Fobs|.
      d Rfree is the Rcryst value calculated on the 5% reflections excluded for refinement.
      The structure of AfAA11A (Fig. 2) shows a typical LPMO fold with a core composed of β-strands that are connected by several loops and a few short helices. The core of the protein consists of two β-sheets formed by antiparallel β-strands, which are arranged in an immunoglobulin-like β-sandwich fold. Similar to AA9s, and unlike AA10s, AfAA11A has an extended region C terminally of the β8-strand, that is, the most C-terminal strand in the β-sandwich. Pairwise alignments with all structures in the Protein Data Bank using the PDBeFold service (
      • Krissinel E.
      • Henrick K.
      Secondary-structure matching (SSM), a new tool for fast protein structure alignment in three dimensions.
      ) at default settings showed that AfAA11A is structurally most similar to the catalytic domain of AoAA11 (Protein Data Bank ID: 4MAH), with a Q score of 0.58 and a Cα-RMSD of 1.59 over 186 residues. These two proteins share 48% sequence identity (Fig. 2A). AfAA11A contains a typical LPMO copper-binding site comprised of the nonmethylated N-terminal histidine and His71 (Figs. 2 and 3). The axial copper coordination positions are occupied by the hydroxyl group of Tyr133 and the methyl side chain of Ala69. This is a common arrangement in LPMOs, including chitin-active AA10s, although the amino acid residues in these axial positions may vary. In particular, bacterial chitin-active AA10 LPMOs have a phenylalanine rather than a tyrosine (Tyr133 in AfAA11A) in the proximal axial coordination position. Figure 3B shows that the copper-binding regions of AfAA11A and AoAA11 are almost identical. The variation in the orientation of His1 is likely because of the absence of a metal ion in AfAA11A, since NMR studies have shown that metal binding affects His1 in particular (
      • Aachmann F.L.
      • Sørlie M.
      • Skjak-Braek G.
      • Eijsink V.G.H.
      • Vaaje-Kolstad G.
      NMR structure of a lytic polysaccharide monooxygenase provides insight into copper binding, protein dynamics, and substrate interactions.
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      • Walton P.H.
      • Aachmann F.L.
      Mechanistic basis of substrate-O2 coupling within a chitin-active lytic polysaccharide monooxygenase: An integrated NMR/EPR study.
      ).
      Figure thumbnail gr2
      Figure 2Pairwise sequence alignment of AfAA11A and AoAA11 and structural topology of AfAA11A. A, structural alignment of AfAA11A and AoAA11 (Protein Data Bank ID: 4MAH) generated with the PyMOL alignment tool at default settings. The alignment file was further processed using the ESPript 3 server and Adobe Illustrator. The three pairs of cysteines forming disulfide bridges are labeled with green numbers. Residues missing in the crystal structure of AoAA11 are highlighted by gray shading. Secondary structure elements are indicated above (AfAA11A) and below (AoAA11) the sequence, as follows: α, alpha helix; η, 310 helix; β, beta-strand; T, turn. B, topology diagram of AfAA11A. C, tertiary structure of AfAA11A, with the same color coding as in panel B; secondary structure elements that are connected by a disulfide bridge (yellow; numbering as in panel A) appear in the same color (red, blue, or green). Panel C also shows the side chains of the two histidines of the copper-binding His brace (H1 and H71), a buried tyrosine beneath the histidine brace (Y133), and a surface-exposed tyrosine (Y25).
      Figure thumbnail gr3
      Figure 3Comparison of chitin-active LPMOs. A, the structures of AfAA11A, AoAA11, and CBP21, colored by secondary structure (helices red, strands yellow) and with additional coloring according to the sequence alignment underneath, showing side chains of residues involved in copper coordination and substrate binding (see text for details). The 11-residue insertion in AfAA11A relative to AoAA11 that includes Tyr25 appears in pink, and residues adjacent to this insertion are labeled. The 14-residue insertion near residue 100 in AoAA11 appears in blue, and residues adjacent to the insertion are labeled in both AA11 structures; note that 11 of these residues (gray shading in the sequence) could not be modeled in the AoAA11 structure. Eight of the nine residues forming an insertion near residue 165 in AoAA11 as well as 11 preceding residues could not be modeled in the AoAA11 structure; this 19-residue region is colored green, and adjacent residues are labeled. B, superposition of AfAA11A (green carbons) and AoAA11 (gray carbons); the picture shows side chains in and near the metal-binding site and a zinc atom bound to AoAA11 (purple sphere). C, a view on the putative substrate-binding surfaces of the three LPMOs shown in panel A, colored by electrostatic surface potential calculated using PyMOL. Note that the AoAA11 structure is incomplete and that the real surface thus may look differently (see main text). The arrows point at a small pocket near the catalytic center that seems to be conserved in chitin-active LPMOs, across LPMO families. LPMO, lytic polysaccharide monooxygenase.
      While the two AA11 enzymes have almost identical catalytic sites, they show considerable differences elsewhere in the structure. Relative to AoAA11, AfAA11A has an insertion spanning residues 16 to 26 (Figs. 2A and 3A), which contains a surface-exposed tyrosine, Tyr25, in a position similar to that of Tyr27 in CBP21 (SmAA10A), a chitin-active AA10 (
      • Vaaje-Kolstad G.
      • Houston D.R.
      • Riemen A.H.
      • Eijsink V.G.H.
      • van Aalten D.M.
      Crystal structure and binding properties of the Serratia marcescens chitin-binding protein CBP21.
      ) (Fig. 3A). Previous studies have shown that this residue is important for chitin binding (
      • Vaaje-Kolstad G.
      • Houston D.R.
      • Riemen A.H.
      • Eijsink V.G.H.
      • van Aalten D.M.
      Crystal structure and binding properties of the Serratia marcescens chitin-binding protein CBP21.
      ,
      • Zeltins A.
      • Schrempf H.
      Specific interaction of the Streptomyces chitin-binding protein Chb1 with α-chitin – the role of individual tryptophan residues.
      ,
      • Bissaro B.
      • Isaksen I.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      • Røhr Å.K.
      How a lytic polysaccharide monooxygenase binds crystalline chitin.
      ). AoAA11 carries two insertions relative to AfAA11A, which both have unknown locations as a major part of both of these is lacking in the AoAA11 crystal structure, as shown in Figure 3A. From the location of these insertions, it seems unlikely that they are located in the same area as the Tyr25-containing insertion in AfAA11A. Next to most of these two insertions, the AoAA11 structure lacked additional residues (151–161) that are resolved in the AfAA11A structure (residues 144–154), showing that these are located far away from the copper-binding site and the substrate-binding surface. Importantly, the insertions in AoAA11 and the additional region that is not visible in the AoAA11 structure contain a tyrosine or other aromatic residues.
      Although some uncertainty exists regarding the surface of AoAA11, it is clear that the two AA11 enzymes have very different substrate-binding surfaces, as illustrated in Figure 3C, which shows that the substrate-binding surfaces of the two enzymes differ in both shape and electrostatic potential. Interestingly, both AA11 enzymes as well as CBP21 contain a small pocket close to the histidine brace, which has been previously associated with chitin-active LPMOs (
      • Forsberg Z.
      • Nelson C.E.
      • Dalhus B.
      • Mekasha S.
      • Loose J.S.
      • Crouch L.I.
      • Røhr Å.K.
      • Gardner J.G.
      • Eijsink V.G.H.
      • Vaaje-Kolstad G.
      Structural and functional analysis of a lytic polysaccharide monooxygenase important for efficient utilization of chitin in Cellvibrio japonicus.
      ,
      • Vaaje-Kolstad G.
      • Bøhle L.A.
      • Gåseidnes S.
      • Dalhus B.
      • Bjøras M.
      • Mathiesen G.
      • Eijsink V.G.H.
      Characterization of the chitinolytic machinery of Enterococcus faecalis V583 and high-resolution structure of its oxidative CBM33 enzyme.
      ). While it has been proposed that this pocket accommodates an acetyl group (
      • Vaaje-Kolstad G.
      • Bøhle L.A.
      • Gåseidnes S.
      • Dalhus B.
      • Bjøras M.
      • Mathiesen G.
      • Eijsink V.G.H.
      Characterization of the chitinolytic machinery of Enterococcus faecalis V583 and high-resolution structure of its oxidative CBM33 enzyme.
      ), recent modeling data suggest that this pocket may give room to O2 or H2O2 in chitin-active LPMOs (
      • Bissaro B.
      • Streit B.
      • Isaksen I.
      • Eijsink V.G.H.
      • Beckham G.T.
      • DuBois J.L.
      • Røhr Å.K.
      Molecular mechanism of the chitinolytic peroxygenase reaction.
      ) as in accordance with suggestions by Hemsworth et al. (
      • Hemsworth G.R.
      • Taylor E.J.
      • Kim R.Q.
      • Gregory R.C.
      • Lewis S.J.
      • Turkenburg J.P.
      • Parkin A.
      • Davies G.J.
      • Walton P.H.
      The copper active site of CBM33 polysaccharide oxygenases.
      ).

      Substrate specificity

      The substrate specificity of AfAA11A was assessed by incubating the LPMO overnight with a variety of substrates in the presence of 1 mM ascorbic acid (AscA). Chromatographic analyses did not reveal any reductant-dependent product formation in reactions with phosphoric acid-swollen cellulose (PASC), Avicel, tamarind xyloglucan, birchwood xylan, beechwood xylan, acetyl glucuronoxylan from aspen, ivory nut mannan, acetylated konjac glucomannan, potato starch, heparin, hyaluronic acid, and chitosan. In contrast, soluble reaction products were observed for the reactions with chitinous substrates (α-chitin from shrimp shell and β-chitin from squid pen), and only if AscA was present (Fig. 4). Both chromatographic and MS analyses of the products generated from α-chitin and β-chitin showed that these were C1-oxidized chito-oligomers with a degree of polymerization 2 to 8 (Fig. 4C) (
      • Mekasha S.
      • Forsberg Z.
      • Dalhus B.
      • Bacik J.P.
      • Choudhary S.
      • Schmidt-Dannert C.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      Structural and functional characterization of a small chitin-active lytic polysaccharide monooxygenase domain of a multi-modular chitinase from Jonesia denitrificans.
      ). The mass spectra showed clusters of signals that are typical for C1-oxidized oligomers, including the diagnostic peak corresponding to the Na+ or K+ salts of aldonic acids. For example, in the degree of polymerization 6 cluster (Fig. 4B), we observed signals corresponding to the Na+ adducts of the lactone (m/z 1257.375) and aldonic acid (m/z 1275.378) forms and of the sodium salt of the aldonic acid (m/z 1297.356) as well as the K+ adducts of the aldonic acid (m/z 1291.343) and of the aldonic acid binding one Na+ and one K+ ion (m/z 1313.315).
      Figure thumbnail gr4
      Figure 4MALDI-TOF MS and chromatographic analysis of product mixtures generated by AfAA11A from chitin. The reactions contained 1 μM LPMO, 1% (A and B) or 0.6% (C) (w/v) substrate, and 1 mM AscA in 50 mM bis–Tris/HCl buffer, pH 6.5, and were incubated at 30 °C with shaking at 1000 rpm for 24 h. A, MALDI-TOF MS spectra showing products generated by AfAA11A from α-chitin or β-chitin with AscA; typical clusters of signals that represent oxidized chito-oligosaccharides of different DP are indicated. The insets show the negative control reactions, without either LPMO (α + AscA; β + AscA) or AscA (α + AfAA11A; β + AfAA11A). B, detailed view and annotation of the DP6 cluster from the reaction with α-chitin shown in panel A. C, HILIC analysis of the reaction products; products were identified using a standard containing a mixture of C1-oxidized chito-oligosaccharides with DP 2 to 6. AscA, ascorbic acid; DP, degree of polymerization; HILIC, hydrophilic interaction liquid chromatography; LPMO, lytic polysaccharide monooxygenase.

      Degradation of chitin in O2-driven reactions

      The ability of AfAA11A to degrade α-chitin and β-chitin under conditions that are commonly used when studying LPMOs (aerobic reactions, 1 mM AscA) was initially tested at temperatures between 30 and 45 °C, to find a temperature giving stable reaction kinetics that could be used in subsequent experiments. The product accumulation over time indicates not only faster LPMO rates but also increased nonlinearity, with increasing temperatures (Fig. 5, A and B). While product formation was linear for the first 6 h at all the three tested temperatures (Fig. 5C), only in reactions run at 30 °C, the product formation rate was stable during the full 24 h of the reaction. Hence, all subsequent reactions in this study were run at 30 °C.
      Figure thumbnail gr5
      Figure 5Activity and binding of AfAA11A on α-chitin and β-chitin. The reactions (AC) contained 1 μM LPMO, 1% (w/v) substrate, and 1 mM AscA in 50 mM bis–Tris/HCl buffer, pH 6.5, and were incubated with shaking at 1000 rpm using three different temperatures, as indicated. Before quantification by HPLC, the solubilized reaction products were treated with chitobiase to convert oxidized oligomers to the oxidized dimer (GlcNAcGlcNAc1A). Panels A and B show progress curves for reactions run at 30, 37, and 45 °C with α-chitin (A) or β-chitin (B). Panel C shows a linear regression of data points for the first 6 h derived from panels A and B. Binding reactions (D) contained 3.0 μM LPMO and 0.2% (w/v) α-chitin (open circles) or β-chitin (filled circles) in 50 mM bis–Tris/HCl buffer, pH 6.5, and were incubated at 30 °C and 1000 rpm. The graph shows the fraction of the protein remaining free in solution. Error bars indicate standard deviations for three independent reactions. AscA, ascorbic acid; LPMO, lytic polysaccharide monooxygenase.
      Figure 5 also shows that AfAA11A is somewhat more active on β-chitin compared with α-chitin, both in terms of the initial rate and the product yield after 24 h. Binding studies showed that AfAA11A binds better to β-chitin compared with α-chitin (Fig. 5D), which could explain the higher activity toward the former substrate. It is worth noting that the progress curves shown in Figure 5 are similar to those typically obtained with chitin-active bacterial AA10 LPMOs when using similar reaction conditions (e.g., (
      • Mutahir Z.
      • Mekasha S.
      • Loose J.S.M.
      • Abbas F.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      • Forsberg Z.
      Characterization and synergistic action of a tetra-modular lytic polysaccharide monooxygenase from Bacillus cereus.
      )). The low catalytic rates that can be estimated from the curves shown in Figure 5, AC (in the range of 0.002–0.01 s−1) are commonly observed for LPMOs in AscA-/O2-driven reactions (
      • Bissaro B.
      • Várnai A.
      • Røhr Å.K.
      • Eijsink V.G.H.
      Oxidoreductases and reactive oxygen species in conversion of lignocellulosic biomass.
      ). Progress curves generated after storing the copper-saturated enzyme for 15 months in 50 mM bis–Tris/HCl, pH 6.5, at 4 °C were similar to those shown in Figure 5, indicating that purified AfAA11A had good storage stability.

      Degradation of chitin in H2O2-driven reactions

      Recent studies have shown that feeding LPMOs with H2O2 may lead to fast turnover of chitin or cellulose (
      • Bissaro B.
      • Røhr Å.K.
      • Muller G.
      • Chylenski P.
      • Skaugen M.
      • Forsberg Z.
      • Horn S.J.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2.
      ,
      • Kuusk S.
      • Bissaro B.
      • Kuusk P.
      • Forsberg Z.
      • Eijsink V.G.H.
      • Sørlie M.
      • Väljamäe P.
      Kinetics of H2O2-driven degradation of chitin by a bacterial lytic polysaccharide monooxygenase.
      ), and one study, describing a novel assay to detect LPMO-generated carboxyl groups in chitin (
      • Wang D.
      • Li J.
      • Wong A.C.Y.
      • Aachmann F.L.
      • Hsieh Y.S.Y.
      A colorimetric assay to rapidly determine the activities of lytic polysaccharide monooxygenases.
      ), has indicated that this may also be the case for members of the AA11 family. Therefore, we analyzed if and how H2O2 could be used to boost the catalytic activity of AfAA11A. For our experiments, we repeatedly added varying amounts of H2O2 to reactions containing the substrate, LPMO, and AscA. Thus, we mimicked a situation where H2O2 is delivered even more gradually, for example, by a H2O2-generating enzyme (
      • Manavalan T.
      • Stepnov A.A.
      • Hegnar O.A.
      • Eijsink V.G.H.
      Sugar oxidoreductases and LPMOs – two sides of the same polysaccharide degradation story?.
      ) or through continuous feeding with a pump, which has shown to be highly effective for cellulose-active LPMOs (
      • Bissaro B.
      • Røhr Å.K.
      • Muller G.
      • Chylenski P.
      • Skaugen M.
      • Forsberg Z.
      • Horn S.J.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2.
      ,
      • Müller G.
      • Chylenski P.
      • Bissaro B.
      • Eijsink V.G.H.
      • Horn S.J.
      The impact of hydrogen peroxide supply on LPMO activity and overall saccharification efficiency of a commercial cellulase cocktail.
      ,
      • Kont R.
      • Bissaro B.
      • Eijsink V.G.H.
      • Väljamäe P.
      Kinetic insights into the peroxygenase activity of cellulose-active lytic polysaccharide monooxygenases (LPMOs).
      ). When H2O2 was added in lower concentrations (20–50 μM H2O2; Fig. 6A), the accumulation of soluble oxidized products correlated well with the amount of H2O2 added, indicating that H2O2 was consumed productively between additions. After the H2O2 feeding phase (Fig. 6A), the LPMO reaction proceeded at the same rate as in the control reaction with only AscA, confirming that H2O2 had been consumed during the feeding phase and that the LPMO was still active. Increasing the amount of H2O2 (80–200 μM per addition; Fig. 6B) gave higher initial product formation rates combined with increasingly rapid cessation of product formation. The observation that the LPMOs become inactivated when supplied with (too) high amounts of H2O2 is in accordance with observations made for LPMOs in the AA9 and AA10 families (
      • Bissaro B.
      • Røhr Å.K.
      • Muller G.
      • Chylenski P.
      • Skaugen M.
      • Forsberg Z.
      • Horn S.J.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2.
      ,
      • Müller G.
      • Chylenski P.
      • Bissaro B.
      • Eijsink V.G.H.
      • Horn S.J.
      The impact of hydrogen peroxide supply on LPMO activity and overall saccharification efficiency of a commercial cellulase cocktail.
      ).
      Figure thumbnail gr6
      Figure 6H2O2-mediated degradation of β-chitin. Reactions contained 1% (w/v) β-chitin, varying amounts of LPMO, and 1 mM AscA in 50 mM bis–Tris/HCl buffer, pH 6.5, and were incubated at 30 °C with shaking at 1000 rpm. Reactions were supplemented periodically with various amounts of H2O2 as indicated in the figure and detailed here. At t = 0, the reactions were supplied with LPMO, AscA, and then H2O2. A, reactions containing 1 μM LPMO were supplied with varying amounts of H2O2 (0, 20, 35, and 50 μM) added every 15 min from t = 0 up to the 3 h mark (vertical dotted line; last addition at 2.75 h). B, reactions containing 1 μM LPMO were supplied with varying amounts of H2O2 (50–200 μM) added every 15 min from t = 0 h to t = 6 h (Note the fewer sampling points within the first 3 h compared with panel A). C, reactions containing varying amounts of the LPMO (0.05, 0.1, 0.2, 0.4, and 1 μM) were supplied with 40 μM H2O2 added every 30 min from t = 0 h to t = 4 h. Error bars indicate standard deviations derived from three independent reactions. No oxidized products were observed in control reactions lacking AscA or the LPMO (not shown). Reaction products were quantified by HPLC after treatment with chitobiase. AscA, ascorbic acid; H2O2, hydrogen peroxide; LPMO, lytic polysaccharide monooxygenase.
      To get an impression of the maximum rate of H2O2 consumption by the LPMO, we ran reactions with reduced LPMO concentrations. The results (Fig. 6C) show that, at LPMO concentrations below 0.2 μM, the amount of LPMO became limiting for the reaction. On the other hand, dose–response effects were minimal at LPMO concentrations of 0.2 μM and higher, suggesting that in all these reactions, H2O2 consumption was complete and that H2O2 was limiting. The apparent initial rates that can be derived from the progress curves for the reactions with 0.05 and 0.1 μM LPMO are in the range of 0.15 to 0.20 s−1, which is 1 to 2 orders of magnitude higher than the apparent rates observed in standard reactions with only AscA and no added H2O2. Of note, the apparent rates of AfAA11A are likely underestimated because a significant fraction of the oxidized sites, perhaps up to 50% (
      • Loose J.S.
      • Forsberg Z.
      • Kracher D.
      • Scheiblbrandner S.
      • Ludwig R.
      • Eijsink V.G.H.
      • Vaaje-Kolstad G.
      Activation of bacterial lytic polysaccharide monooxygenases with cellobiose dehydrogenase.
      ), may remain in the solid residue after saccharification. It is also worth noting that the linear progress curve for the reaction with 0.2 μM AfAA11A in Figure 6C, where the level of soluble product reaches 250 μM after 4 h, implies that, in this experimental setting, one LPMO molecule can turnover at least some 1250 molecules of H2O2 without becoming inactivated.

      Synergy with a chitinase

      The ability of AfAA11A to boost chitinase efficiency was assessed by monitoring the degradation of α-chitin and β-chitin by a combination of an endochitinase, chitinase C from SmChiC (ChiC), and AfAA11A in a 1:1 M ratio (Fig. 7). The results show that AfAA11A promotes the activity of SmChiC on both α-chitin and β-chitin. When acting individually on α-chitin for 24 h (Fig. 7A), the proteins were able to solubilize 1.7 and 2.6 mM GlcNAc of the substrate, respectively, while when combined, 12.6 mM GlcNAc was produced. In the reactions with α-chitin and SmChiC only, product formation stopped after some 9 h, whereas linear product formation was achieved for 24 h in the reaction that also contained the LPMO.
      Figure thumbnail gr7
      Figure 7Synergistic action of AfAA11A and a chitinase in chitin degradation. The figure shows chitin solubilization over time in various reactions with α-chitin (A) or β-chitin (B). All reaction mixtures contained 1 μM LPMO and/or 1 μM ChiC, and approximately 10 g/l α-chitin or 20 g/l β-chitin in 50 mM bis–Tris/HCl buffer, pH 6.5, and were incubated at 30 °C with shaking at 1000 rpm. For reactions containing the LPMO, 1 mM AscA was included. Before quantification, reaction products were treated with chitobiase to convert all soluble products to a mixture of GlcNAc, which was quantified, and minor amounts of GlcNAcGlcNAc1A. The vertical dotted lines indicate the time point for addition of AfAA11A in the reactions labeled “ChiC + AfAA11A (6 h).” Error bars indicate standard deviations for three independent reactions. AscA, ascorbic acid; ChiC, SmChiC; LPMO, lytic polysaccharide monooxygenase.
      Synergistic effects were also observed in reactions with β-chitin (Fig. 7B). Compared with the reactions with α-chitin, these effects were smaller, which may be due to the fact that the chitinase alone was much more efficient on β-chitin (Fig. 7B) than on α-chitin (Fig. 7A). The initial rate of product formation by SmChiC was little affected by the addition of AfAA11A. On the other hand, in the reaction where SmChiC was supplemented with the LPMO, the high initial rate (as observed with SmChiC alone) was maintained for a longer time, and, consequently, higher product levels were reached earlier on in the reaction. Final saccharification yields after 24 h were also affected by the LPMO. While SmChiC and AfAA11A alone were able to produce 54.9 and 2.7 mM GlcNAc, respectively, 84.6 mM GlcNAc was produced in the reaction containing both enzymes.
      To highlight the impact of the LPMO in chitin depolymerization, and to verify whether the cessation of product formation in reactions with SmChiC was caused by enzyme inactivation, denaturation, or depletion of accessible substrate, reactions were carried out in which the LPMO was added after 6 h, instead of at the start of the reaction. This experiment showed that chitin solubilization by SmChiC was enhanced upon addition of the LPMO (Fig. 7, A and B) and that, in these reactions, product formation continued for the remaining reaction period. Thus, the gradual cessation of product formation in reactions with only SmChiC is due to the enzyme not being able to access the remaining substrate and not to enzyme inactivation.

      Discussion

      The crystal structure of AfAA11A is only the second crystal structure of a member of the widely spread family of fungal AA11 LPMOs and the first one that shows all amino acids. The AfAA11A structure lacks a metal ion, which is remarkable considering the reported high metal affinities of these enzymes (
      • Quinlan R.J.
      • Sweeney M.D.
      • Lo Leggio L.
      • Otten H.
      • Poulsen J.C.N.
      • Johansen K.S.
      • Krogh K.B.R.M.
      • Jorgensen C.I.
      • Tovborg M.
      • Anthonsen A.
      • Tryfona T.
      • Walter C.P.
      • Dupree P.
      • Xu F.
      • Davies G.J.
      • et al.
      Insights into the oxidative degradation of cellulose by a copper metalloenzyme that exploits biomass components.
      ,
      • Aachmann F.L.
      • Sørlie M.
      • Skjak-Braek G.
      • Eijsink V.G.H.
      • Vaaje-Kolstad G.
      NMR structure of a lytic polysaccharide monooxygenase provides insight into copper binding, protein dynamics, and substrate interactions.
      ) but not uncommon (e.g., (
      • Vaaje-Kolstad G.
      • Houston D.R.
      • Riemen A.H.
      • Eijsink V.G.H.
      • van Aalten D.M.
      Crystal structure and binding properties of the Serratia marcescens chitin-binding protein CBP21.
      )). Nevertheless, the catalytic centers of AfAA11A and AoAA11, which has a zinc ion bound, look similar (Fig. 3B). Comparison of the structures of AfAA11A and AoAA11 beyond the catalytic centers is complicated because the latter structure lacks two regions comprising 30 residues in total. Residues 144 to 154 in the structure of AfAA11A provide structural information for residues 151 to 161 lacking in the structure of AoAA11. It is somewhat remarkable that these residues form an α-helix (α1; Fig. 2), since the fact that these residues could not be modeled in the AoAA11 structure would suggest that they are part of a flexible loop-like structure.
      Even when taking into account the lacking information for AoAA11, it seems safe to conclude from the comparison of the two structures (Fig. 3) that the two enzymes have different substrate-binding surfaces. Because of the Tyr25-containing insertion that is unique for AfAA11A, the substrate-binding surface of this enzyme shows features that are characteristic for chitin-active AA10 LPMOs, such as CBP21, in particular the presence of exposed Tyr25 (Fig. 3). Experimental (
      • Vaaje-Kolstad G.
      • Houston D.R.
      • Riemen A.H.
      • Eijsink V.G.H.
      • van Aalten D.M.
      Crystal structure and binding properties of the Serratia marcescens chitin-binding protein CBP21.
      ) and modeling (
      • Bissaro B.
      • Isaksen I.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      • Røhr Å.K.
      How a lytic polysaccharide monooxygenase binds crystalline chitin.
      ) studies have shown that a tyrosine in this position is important for binding of crystalline chitin. As alluded to above, it is not possible to envisage a similar structural arrangement in AoAA11 or in previously characterized FfAA11 (
      • Wang D.
      • Li J.
      • Salazar-Alvarez G.
      • McKee L.S.
      • Srivastava V.
      • Sellberg J.A.
      • Bulone V.
      • Hsieh Y.S.Y.
      Production of functionalised chitins assisted by fungal lytic polysaccharide monooxygenase.
      ), which both lack this insertion. Interestingly, these structural differences seem to correlate with functional properties. On the one hand, we show here that the catalytic activity of AfAA11A and its ability to boost enzymatic chitin conversion are similar to what has been observed for bacterial chitin-active AA10 LPMOs. On the other hand, a recent study has shown that another AA11 LPMO from A. fumigatus, AfAA11B, which is a close homolog of AoAA11, with similar insertions and deletions relative to AfAA11A, has negligible activity toward chitin (
      • Rieder L.
      • Petrović D.
      • Väljamäe P.
      • Eijsink V.G.H.
      • Sørlie M.
      Kinetic characterization of a putatively chitin-active LPMO reveals a preference for soluble substrates and absence of monooxygenase activity.
      ). Of note, AfAA11B shows high activity toward soluble chito-oligomers; the biological role of this activity remains enigmatic (
      • Rieder L.
      • Petrović D.
      • Väljamäe P.
      • Eijsink V.G.H.
      • Sørlie M.
      Kinetic characterization of a putatively chitin-active LPMO reveals a preference for soluble substrates and absence of monooxygenase activity.
      ).
      Our data show that AfAA11A has C1-oxidizing activity leading to production of lactone products that are in equilibrium with their aldonic acid forms. It is worth mentioning that the MALDI-TOF MS analyses suggested the presence of considerable amounts of lactones (M-2 Da) in reactions with both α-chitin and β-chitin, which could represent C4-oxidized products (since C1-oxidized lactones are expected to be hydrated to a larger extent (
      • Eijsink V.G.H.
      • Petrović D.
      • Forsberg Z.
      • Mekasha S.
      • Røhr Å.K.
      • Várnai A.
      • Bissaro B.
      • Vaaje-Kolstad G.
      On the functional characterization of lytic polysaccharide monooxygenases (LPMOs).
      )). While this observation, which is similar to observations by Wang et al. (
      • Wang D.
      • Li J.
      • Salazar-Alvarez G.
      • McKee L.S.
      • Srivastava V.
      • Sellberg J.A.
      • Bulone V.
      • Hsieh Y.S.Y.
      Production of functionalised chitins assisted by fungal lytic polysaccharide monooxygenase.
      ) for FfAA11, could indicate the presence of C4-oxidized reaction products, we did not detect unusual, that is, possibly C4-oxidized products in either hydrophilic interaction liquid chromatography (Fig. 5C) or high-performance anion exchange chromatography with pulsed amperometric detection (not shown). Compared with α-chitin, reactions with β-chitin showed both better binding at 30 °C and more product formation at all three tested reaction temperatures (Fig. 5). Other chitin-active LPMOs show similar differences regarding binding to and activity on these two chitin allomorphs (
      • Mekasha S.
      • Forsberg Z.
      • Dalhus B.
      • Bacik J.P.
      • Choudhary S.
      • Schmidt-Dannert C.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      Structural and functional characterization of a small chitin-active lytic polysaccharide monooxygenase domain of a multi-modular chitinase from Jonesia denitrificans.
      ,
      • Nakagawa Y.S.
      • Kudo M.
      • Loose J.S.
      • Ishikawa T.
      • Totani K.
      • Eijsink V.G.H.
      • Vaaje-Kolstad G.
      A small lytic polysaccharide monooxygenase from Streptomyces griseus targeting α- and β-chitin.
      ).
      Since the discovery that LPMOs can use H2O2 as a cosubstrate in 2017 (
      • Bissaro B.
      • Røhr Å.K.
      • Muller G.
      • Chylenski P.
      • Skaugen M.
      • Forsberg Z.
      • Horn S.J.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2.
      ), multiple studies have demonstrated that addition of H2O2 to LPMO reactions leads to increased catalytic rates (
      • Bissaro B.
      • Røhr Å.K.
      • Muller G.
      • Chylenski P.
      • Skaugen M.
      • Forsberg Z.
      • Horn S.J.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2.
      ,
      • Kuusk S.
      • Bissaro B.
      • Kuusk P.
      • Forsberg Z.
      • Eijsink V.G.H.
      • Sørlie M.
      • Väljamäe P.
      Kinetics of H2O2-driven degradation of chitin by a bacterial lytic polysaccharide monooxygenase.
      ,
      • Hedison T.M.
      • Breslmayr E.
      • Shanmugam M.
      • Karnpakdee K.
      • Heyes D.J.
      • Green A.P.
      • Ludwig R.
      • Scrutton N.S.
      • Kracher D.
      Insights into the H2O2-driven catalytic mechanism of fungal lytic polysaccharide monooxygenases.
      ,
      • Wang D.
      • Li J.
      • Wong A.C.Y.
      • Aachmann F.L.
      • Hsieh Y.S.Y.
      A colorimetric assay to rapidly determine the activities of lytic polysaccharide monooxygenases.
      ,
      • Müller G.
      • Chylenski P.
      • Bissaro B.
      • Eijsink V.G.H.
      • Horn S.J.
      The impact of hydrogen peroxide supply on LPMO activity and overall saccharification efficiency of a commercial cellulase cocktail.
      ,
      • Hegnar O.A.
      • Petrović D.M.
      • Bissaro B.
      • Alfredsen G.
      • Várnai A.
      • Eijsink V.G.H.
      pH-dependent relationship between catalytic activity and hydrogen peroxide production shown via characterization of a lytic polysaccharide monooxygenase from Gloeophyllum trabeum.
      ). Indeed, and in accordance with previous observations for FfAA11 (
      • Wang D.
      • Li J.
      • Wong A.C.Y.
      • Aachmann F.L.
      • Hsieh Y.S.Y.
      A colorimetric assay to rapidly determine the activities of lytic polysaccharide monooxygenases.
      ) and AfAA11B (
      • Rieder L.
      • Petrović D.
      • Väljamäe P.
      • Eijsink V.G.H.
      • Sørlie M.
      Kinetic characterization of a putatively chitin-active LPMO reveals a preference for soluble substrates and absence of monooxygenase activity.
      ), we observed increased catalytic rate upon adding H2O2 to reactions with AfAA11A. The apparent catalytic rates of AfAA11A were in the range of 0.003 to 0.012 s−1 for the O2-driven reaction and in the range of 0.1 to 0.2 s−1 for the H2O2-driven reaction. An earlier study on FfAA11A, using the similar reaction conditions (10 g/l α-chitin, 2 μM LPMO, and 1 mM AscA) reported apparent rates of 0.5 min−1 (0.01 s−1) under standard aerobic conditions without H2O2 and a 16-fold increase (to 0.11 s−1) when 100 μM H2O2 was supplied (
      • Wang D.
      • Li J.
      • Wong A.C.Y.
      • Aachmann F.L.
      • Hsieh Y.S.Y.
      A colorimetric assay to rapidly determine the activities of lytic polysaccharide monooxygenases.
      ). Of note, some claim that the so-called “O2-driven” reaction under standard aerobic conditions may be limited by the in situ generation of H2O2, which is needed to fuel the LPMO peroxygenase reaction (see e.g., (
      • Bissaro B.
      • Røhr Å.K.
      • Muller G.
      • Chylenski P.
      • Skaugen M.
      • Forsberg Z.
      • Horn S.J.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2.
      ,
      • Stepnov A.A.
      • Forsberg Z.
      • Sørlie M.
      • Nguyen G.-S.
      • Wentzel A.
      • Røhr Å.K.
      • Eijsink V.G.H.
      Unraveling the roles of the reductant and free copper ions in LPMO kinetics.
      ), and see (
      • Hangasky J.A.
      • Iavarone A.T.
      • Marletta M.A.
      Reactivity of O2 versus H2O2 with polysaccharide monooxygenases.
      ) for another opinion). The apparent rates for the peroxygenase reaction are much lower than a kcat value reported for CBP21 acting on chitin (6.7 s−1; (
      • Kuusk S.
      • Bissaro B.
      • Kuusk P.
      • Forsberg Z.
      • Eijsink V.G.H.
      • Sørlie M.
      • Väljamäe P.
      Kinetics of H2O2-driven degradation of chitin by a bacterial lytic polysaccharide monooxygenase.
      )). This may reflect a true difference in enzyme efficiency, but it is important to note that the peroxygenase rate of AfAA11A is underestimated because of noncontinuous product monitoring, which may hide possibly much higher rates that are achieved right after each addition of H2O2. Additional kinetic studies are needed to determine the true maximum rate of AfAA11A.
      The experiments depicted in Figure 6 show that the AfAA11A peroxygenase reaction may be limited by H2O2 (Fig. 6A) or by the LPMO concentration (Fig. 6C), depending on how the reaction is set up. The H2O2-limited reactions in Figure 6A show a strong correlation between the amount of added H2O2 and the amount of soluble oxidized products. In each of the three reactions with added H2O2, the level of oxidized products at the end of the H2O2-feeding period (and corrected for product levels generated in the control reaction with reductant only) correspond to some 60 to 65% of the total amount of added H2O2. It is well known from the literature that a considerable portion, in some cases up to 50%, of the oxidized sites introduced by LPMOs remain on the crystalline substrate (
      • Courtade G.
      • Forsberg Z.
      • Heggset E.B.
      • Eijsink V.G.H.
      • Aachmann F.L.
      The carbohydrate-binding module and linker of a modular lytic polysaccharide monooxygenase promote localized cellulose oxidation.
      ), which may explain the discrepancy between the amount of added H2O2 and the amount of soluble oxidized products.
      As expected for LPMOs (and other enzymes with active redox centers), excessive H2O2 concentrations lead to enzyme inactivation of LPMOs (
      • Bissaro B.
      • Røhr Å.K.
      • Muller G.
      • Chylenski P.
      • Skaugen M.
      • Forsberg Z.
      • Horn S.J.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2.
      ,
      • Petrović D.M.
      • Bissaro B.
      • Chylenski P.
      • Skaugen M.
      • Sørlie M.
      • Jensen M.S.
      • Aachmann F.L.
      • Courtade G.
      • Várnai A.
      • Eijsink V.G.H.
      Methylation of the N-terminal histidine protects a lytic polysaccharide monooxygenase from auto-oxidative inactivation.
      ). Figure 6B shows that higher H2O2 concentrations lead to higher initial enzyme rates and more rapid enzyme inactivation, as has been demonstrated for multiple LPMOs, including FfAA11 (
      • Wang D.
      • Li J.
      • Wong A.C.Y.
      • Aachmann F.L.
      • Hsieh Y.S.Y.
      A colorimetric assay to rapidly determine the activities of lytic polysaccharide monooxygenases.
      ). Importantly, at lower H2O2 concentrations, the enzyme seems stable, and we show that one enzyme molecule on average catalyzes at least 1250 reactions (note that this by no means is a maximum or an optimized value). This value compares well with previously published values of ≥300 (for AfAA11B acting on soluble substrate, (
      • Rieder L.
      • Petrović D.
      • Väljamäe P.
      • Eijsink V.G.H.
      • Sørlie M.
      Kinetic characterization of a putatively chitin-active LPMO reveals a preference for soluble substrates and absence of monooxygenase activity.
      )) and ≥1500 for the LPMO in a commercial cellulase cocktail (
      • Bissaro B.
      • Røhr Å.K.
      • Muller G.
      • Chylenski P.
      • Skaugen M.
      • Forsberg Z.
      • Horn S.J.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2.
      ).
      Although the progress curves of Figure 5 (i.e., activity of the LPMO only) showed higher activity of AfAA11A on β-chitin compared with α-chitin, the boosting effect of the LPMO on chitinase activity was more pronounced for α-chitin (Fig. 7). α-chitin is more crystalline and resilient to enzymatic degradation than β-chitin (
      • Kaya M.
      • Mujtaba M.
      • Ehrlich H.
      • Salaberria A.M.
      • Baran T.
      • Amemiya C.T.
      • Galli R.
      • Akyuz L.
      • Sargin I.
      • Labidi J.
      On chemistry of γ-chitin.
      ), and it is thus conceivable that the addition of an enzyme specialized in acting on crystalline material, that is, the LPMO, has a larger effect in reactions with α-chitin. Indeed, previous studies have shown that the extent by which an LPMO can speed up a chitinase reaction depends on substrate crystallinity (
      • Nakagawa Y.S.
      • Eijsink V.G.H.
      • Totani K.
      • Vaaje-Kolstad G.
      Conversion of α-chitin substrates with varying particle size and crystallinity reveals substrate preferences of the chitinases and lytic polysaccharide monooxygenase of Serratia marcescens.
      ).
      In conclusion, we present the first structure of an AA11 LPMO in which all amino acids are included and show that there is considerable structural variation within the AA11 family that is correlated to functional variation. It is worth noting that the disclosed variation within the AA11 family really is remarkable. First, the structural differences in the substrate-binding surfaces seem larger than the differences seen within other LPMO families. Second, while AfAA11A degrades chitin with commonly observed reaction rates and yields, AfAA11B does not. We also show that AfAA11A can efficiently utilize H2O2 as a cosubstrate and is not inactivated by acting in the “peroxygenase” mode, as long as H2O2 levels are not too high. AfAA11A enhanced the solubilization of β-chitin and α-chitin by SmChiC, suggesting that this LPMO may have a role in fungal chitin degradation and may be of use in the enzymatic processing of chitin-rich biomass.

      Experimental procedures

      Reagents and substrates

      All common reagents used in this study were supplied by Sigma–Aldrich and Merck Millipore, whereas Bacto Peptone and Bacto Yeast Extract were supplied by BD Biosciences, and acetonitrile was supplied by VWR. Alkaline- and acid- pretreated shrimp shell α-chitin originating from Pandalus borealis was purchased from Chitinor AS and ball-milled using a Planetary Ball Mill PM100 (Retsch) equipped with a stainless-steel container and zirconium dioxide beads. Milling was performed at 450 rpm in three 5-min rounds with 2-min intervals, yielding a final particle size of ∼0.2 mm. β-chitin originating from squid pen (batch: 20140101) with an average particle size of 0.8 mm was purchased from France Chitine.
      Ivory nut mannan, acetylated konjac glucomannan, beechwood xylan, and tamarind xyloglucan were purchased from Megazyme. Birchwood xylan, Avicel, shrimp shell chitosan with approximate mass of 190 to 375 kDa and ≥75% deacetylation, and hyaluronic acid with approximate mass of 0.6 to 1.1 MDa were purchased from Sigma–Aldrich. Potato starch and heparin from pig were purchased from Merck Millipore. Acetyl glucuronoxylan from aspen was produced in-house as described earlier (
      • Biely P.
      • Cziszárová M.
      • Uhliariková I.
      • Agger J.W.
      • Li X.L.
      • Eijsink V.G.H.
      • Westereng B.
      Mode of action of acetylxylan esterases on acetyl glucuronoxylan and acetylated oligosaccharides generated by a GH10 endoxylanase.
      ). PASC was produced in-house from Avicel, as previously described (
      • Wood T.M.
      Preparation of crystalline, amorphous, and dyed cellulase substrates.
      ). Tetra-N-acetylchitotetraose, penta-N-acetylchitopentaose, and hexa-N-acetylchitohexaose were purchased from Megazyme. Chitinase SmChi18C, chitobiase SmGH20 (GenBank ID, AAB03808.1), and endo-β-N-acetylglucosaminidase EfEndo18A were recombinantly produced and purified as described previously (
      • Bohle L.A.
      • Mathiesen G.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      An endo-ss-N-acetylglucosaminidase from Enterococcus faecalis V583 responsible for the hydrolysis of high-mannose and hybrid-type N-linked glycans.
      ,
      • Synstad B.
      • Vaaje-Kolstad G.
      • Cederkvist F.H.
      • Saua S.F.
      • Horn S.J.
      • Eijsink V.G.H.
      • Sørlie M.
      Expression and characterization of endochitinase C from Serratia marcescens BJL200 and its purification by a one-step general chitinase purification method.
      ,
      • Loose J.S.M.
      • Forsberg Z.
      • Fraaije M.W.
      • Eijsink V.G.H.
      • Vaaje-Kolstad G.
      A rapid quantitative activity assay shows that the Vibrio cholerae colonization factor GbpA is an active lytic polysaccharide monooxygenase.
      ).
      N-acetyl-d-glucosamine (GlcNAc) with a purity of >95% used for producing standard curves for quantitative product analysis was obtained from Megazyme. Standards of C1-oxidized chito-oligosaccharides were produced in-house by incubating N-acetyl-chito-oligosaccharides (Megazyme; 95% purity) with Fusarium graminearum chito-oligosaccharide oxidase (FgChitO) as previously described (
      • Loose J.S.M.
      • Forsberg Z.
      • Fraaije M.W.
      • Eijsink V.G.H.
      • Vaaje-Kolstad G.
      A rapid quantitative activity assay shows that the Vibrio cholerae colonization factor GbpA is an active lytic polysaccharide monooxygenase.
      ,
      • Heuts D.P.
      • Winter R.T.
      • Damsma G.E.
      • Janssen D.B.
      • Fraaije M.W.
      The role of double covalent flavin binding in chito-oligosaccharide oxidase from Fusarium graminearum.
      ).

      Protein production and purification

      The AfAA11A protein was produced through heterologous expression using a P. pastoris yeast expression system (PichiaPink Strain 4; Thermo Fisher Scientific). A synthetic gene encoding AfAA11A (UniProt ID: Q4WF00; gene ID: AFUA_3G03950) including its native signal peptide was codon-optimized for P. pastoris and synthesized by GenScript and inserted behind the GAP promoter and a P. pastoris-specific Kozak sequence in the pPink-GAP vector by restriction cloning with EcoRI and Acc65I (New England BioLabs, Inc). The generated plasmid was linearized with AflII (New England BioLabs, Inc) and transformed into electrocompetent P. pastoris cells, after which LPMO-producing transformants were screened, selected, and stored as previously described (
      • Várnai A.
      • Tang C.
      • Bengtsson O.
      • Atterton A.
      • Mathiesen G.
      • Eijsink V.G.H.
      Expression of endoglucanases in Pichia pastoris under control of the GAP promoter.
      ).
      For protein production, a single colony from a yeast extract–peptone–dextrose agar plate was used to inoculate 50 ml of buffered glycerol complex medium (Invitrogen) in a 250-ml Erlenmeyer flask with vent cap, which was then incubated overnight at 29 °C and 200 rpm. The overnight culture was transferred to 400 ml of fresh buffered glycerol complex medium, followed by incubation overnight in a 2-l baffled shake flask with vent cap. After 24 h, the culture was supplemented with 1% (v/v) glycerol. After 48 h, the culture supernatant containing the LPMO was separated from the yeast cells by centrifugation at 10,000g for 20 min at 4 °C and subsequentially filtered through a 0.45 μm polyether sulfone (PES) membrane (VWR). The presence of the LPMO was confirmed by SDS-PAGE gel analysis using Mini PROTEAN TGX Stain-Free precast gels, and fluorescence was recorded with a Gel Doc EZ Imager using the Image Lab, version 6.0.0 Standard Edition software (Bio-Rad). The culture supernatant was concentrated approximately fivefold with concomitant buffer exchange to 50 mM bis–Tris/HCl buffer, pH 6.5, using a Vivaflow 200 tangential crossflow concentrator equipped with a 10,000 molecular weight cutoff (MWCO) PES membrane (Sartorius Stedim Biotech).
      For hydrophobic interaction chromatography, the concentrated culture supernatant was supplemented with ammonium sulphate to a final concentration of 2 M and centrifuged for 15 min at 10,000g and 4 °C to remove precipitation. The supernatant was then applied onto a 5 ml HiTrap Phenyl FF (HS) column (GE Healthcare Lifesciences), mounted on an ÄKTA Prime Plus chromatography system and equilibrated in 50 mM bis–Tris/HCl buffer (pH 6.5) containing 2 M ammonium sulphate. Proteins were eluted by applying a 35 ml linear gradient toward 50 mM bis–Tris/HCl buffer, pH 6.5. Fractions containing AfAA11A were identified by SDS-PAGE, pooled, and concentrated using a Vivaspin 20 Centrifugal Concentrator 10,000 MWCO PES (Sartorius Stedim Biotech). The recombinant protein was further purified by size-exclusion chromatography using a HiLoad 16/600 Superdex 75 pg column (GE Healthcare) mounted on an ÄKTA purifier chromatography system (GE Healthcare), using 50 mM bis–Tris/HCl, pH 6.5, 150 mM NaCl as eluent, and a flow rate of 1 ml·min−1. Fractions containing pure LPMO were identified by SDS-PAGE and pooled, followed by concentration and buffer exchange to 50 mM bis–Tris/HCl buffer, pH 6.5, using a Vivaspin 20 Centrifugal Concentrator 10,000 MWCO PES.
      To saturate the LPMO with Cu(II), the purified protein was incubated with 10-fold molar excess of CuSO4 at 4 °C for 4 h, then desalted on a Vivaspin 20 Centrifugal Concentrator 3000 MWCO PES by five cycles of concentrating and rediluting the protein solution in 50 mM bis–Tris/HCl buffer, pH 6.5 (ca. 10,000-fold dilution). The final protein solution was sterilized by filtration through a 0.22 μm Millex-GV syringe filter (Merck Millipore) and stored at 4 °C.
      Protein concentrations of solutions containing pure AfAA11A were determined based on absorbance at 280 nm with an Eppendorf Biophotometer D30 (Eppendorf), using the theoretical extinction coefficient of 38,765 M−1 cm−1, calculated with the ExPASy ProtParam tool (
      • Wilkins M.R.
      • Gasteiger E.
      • Bairoch A.
      • Sanchez J.C.
      • Williams K.L.
      • Appel R.D.
      • Hochstrasser D.F.
      Protein identification and analysis tools in the ExPASy server.
      ). The mature protein consists of 201 amino acid residues with a calculated molecular mass of 21,750 Da and a theoretical pI of 4.76 (
      • Wilkins M.R.
      • Gasteiger E.
      • Bairoch A.
      • Sanchez J.C.
      • Williams K.L.
      • Appel R.D.
      • Hochstrasser D.F.
      Protein identification and analysis tools in the ExPASy server.
      ).

      Protein crystallization

      N-linked glycosylations were removed from a batch of purified and copper-saturated AfAA11A by treating the LPMO (140 μM) with an in-house–produced recombinant endo-β-N-acetylglucosaminidase (EfEndo18A, 3.4 μM; (
      • Bohle L.A.
      • Mathiesen G.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      An endo-ss-N-acetylglucosaminidase from Enterococcus faecalis V583 responsible for the hydrolysis of high-mannose and hybrid-type N-linked glycans.
      )) in 50 mM sodium acetate buffer (pH 6.0). The reaction (with 5 ml total volume) was incubated first at 30 °C for 1 h and then at 4 °C overnight. Deglycosylation was verified by SDS-PAGE, and the EfEndo18A protein was removed by hydrophobic interaction chromatography, as described previously. Fractions containing the LPMO were pooled and concentrated, and the buffer was exchanged to 50 mM bis–Tris/HCl buffer (pH 6.5) using a Vivaspin 20 Centrifugal Concentrator 10,000 MWCO PES. The resulting protein was saturated with Cu(II) and subsequentially desalted, filter sterilized, and stored according to the protocol described previously. Deglycosylated AfAA11A was successfully crystallized using conditions 1 to 15 from the commercial crystal screening kit JCSG-plus MD 1-37 Box 1 (Molecular Dimensions) in a hanging drop setup using a 24-well VDX plate with sealant and 18 mm circle glass cover slides (Hampton Research). The mother liquor was made in-house and consisted of 0.1 M bicine (pH 9.0) and 20% (w/v) PEG 6000. The best crystals were produced in droplets obtained by mixing 0.5 μl of 20 mg·ml−1 de-N-glycosylated AfAA11A with 1 μl mother liquor and having 200 μl mother liquor in the well. The crystallization setups were stored in the dark at room temperature, and protein crystals were observed after approximately 3.5 months.

      X-ray crystallography

      Protein crystals were soaked in a cryosolution containing mother-liquor with 35% glucose (w/v) before they were flash frozen in liquid nitrogen. Diffraction data were collected at the BioMAX beamline at MAX IV. Datasets were processed by XDS (
      • Kabsch W.
      XDS.
      ) and scaled by AIMLES (
      • Evans P.R.
      • Murshudov G.N.
      How good are my data and what is the resolution?.
      ). The CCP4i package was used to solve the structure by molecular replacement (Phaser) (
      • McCoy A.J.
      • Grosse-Kunstleve R.W.
      • Adams P.D.
      • Winn M.D.
      • Storoni L.C.
      • Read R.J.
      Phaser crystallographic software.
      ), and the structure was refined using REFMAC (
      • Murshudov G.N.
      • Skubak P.
      • Lebedev A.A.
      • Pannu N.S.
      • Steiner R.A.
      • Nicholls R.A.
      • Winn M.D.
      • Long F.
      • Vagin A.A.
      REFMAC5 for the refinement of macromolecular crystal structures.
      ). Model manipulations were carried out using Coot (
      • Emsley P.
      • Lohkamp B.
      • Scott W.G.
      • Cowtan K.
      Features and development of COOT.
      ), and molecular graphics was generated using PyMOL Molecular Graphics System, version 2.0 (Schrödinger, LLC).

      Substrate specificity

      The activity of AfAA11A was screened toward the following substrates: PASC, Avicel, tamarind xyloglucan, birchwood xylan, beechwood xylan, acetyl glucuronoxylan from aspen, ivory nut mannan, acetylated konjac glucomannan, potato starch, heparin, hyaluronic acid, chitosan from shrimp shell, α-chitin from shrimp shell, and β-chitin from squid pen. Reaction mixtures were set up in 100 μl total volume, contained 1 μM LPMO, 0.2 to 0.6% (w/v) substrate, and 1 mM AscA in 50 mM bis–Tris/HCl buffer, pH 6.5, and were incubated overnight at 30 °C and 1000 rpm in an Eppendorf Thermomixer C (Eppendorf). In negative control reactions, AscA or the LPMO was substituted with Milli-Q water. At the end of the reaction, insoluble substrates were removed by filtration using a 0.2 μm PES 96-well filter plate (Millipore) operated in a vacuum manifold, and the soluble fraction was subsequently analyzed as outlined later.

      Chitin degradation experiments

      In general, reactions, with a total volume of 300 μl, were set up with 1% (w/v) substrate and 1 μM LPMO and 1 mM AscA in 50 mM bis–Tris/HCl buffer, pH 6.5, and were incubated at 30 °C and 1000 rpm in thermomixers (Eppendorf Thermomixer C; Eppendorf) in standard aerobic conditions for up to 24 h. Samples (30 μl) were taken periodically, and the reaction was terminated by removing the insoluble substrate by filtration using a 0.2 μm PES 96-well filter plate (Millipore) operated in a vacuum manifold. All reactions were performed in triplicates.
      When assessing synergy between chitinases and the LPMO, reactions were set up with various combinations of 1 μM LPMO (AfAA11A) and 1 μM SmChi18C, a nonprocessive endochitinase from S. marcescens (
      • Synstad B.
      • Vaaje-Kolstad G.
      • Cederkvist F.H.
      • Saua S.F.
      • Horn S.J.
      • Eijsink V.G.H.
      • Sørlie M.
      Expression and characterization of endochitinase C from Serratia marcescens BJL200 and its purification by a one-step general chitinase purification method.
      ). Reactions containing LPMO were supplemented with 1 mM AscA at t = 0. When assessing the effect of H2O2 on β-chitin degradation by AfAA11A, AscA (1 mM) was added at t = 0, after which 30 μl of an appropriate aqueous H2O2 solution was added to reach 20, 35, 40, 50, 80, 120, or 200 μM H2O2 and a total reaction volume of 300 μl. Reactions were sampled regularly and at the same time supplied with a new portion of H2O2. Sample volumes (30 μl) were equivalent to H2O2 addition volumes (30 μl) in order to maintain the reaction volume at 300 μl. Dilution in the amounts of products formed because of sampling and addition of reactant were taken into account when calculating product levels.
      All samples were incubated with 1 μM chitobiase (SmGH20) at 37 °C in a static incubator to convert the solubilized oligomers to a mixture of nonoxidized monosugars (GlcNAc) and C1-oxidized dimers (N,N′-diacetylchitobionic acid, GlcNAcGlcNAc1A), as described previously (
      • Loose J.S.M.
      • Forsberg Z.
      • Fraaije M.W.
      • Eijsink V.G.H.
      • Vaaje-Kolstad G.
      A rapid quantitative activity assay shows that the Vibrio cholerae colonization factor GbpA is an active lytic polysaccharide monooxygenase.
      ).

      Substrate binding

      To assess binding to chitin, 3.0 μM AfAA11A was incubated with 0.2% (w/v) α-chitin or β-chitin in 50 mM bis–Tris/HCl buffer, pH 6.5, at 30 °C and 1000 rpm. At different time points, the insoluble substrate was removed by filtration using a 96-well 0.2 μm PES filter plate (Millipore) installed in a vacuum manifold. The amount of LPMO remaining in solution was visualized on Mini PROTEAN TGX Stain-Free SDS-PAGE gels (Bio-Rad) with a Gel Doc EZ Imager (Bio-Rad) based on fluorescence signals and quantified with the Image Lab software, version 6.0.0. Solutions with known concentrations of AfAA11A were used as standard.

      Analysis and quantification of reaction products

      Product formation in reactions with cellulosic and hemicellulosic substrates was assessed by high-performance anion exchange chromatography with pulsed amperometric detection using an ICS5000 system (Dionex) equipped with CarboPac PA1 analytical (2 × 250 mm) and guard (2 × 50 mm) columns, using a 50 min gradient protocol as previously described (
      • Westereng B.
      • Agger J.W.
      • Horn S.J.
      • Vaaje-Kolstad G.
      • Aachmann F.L.
      • Stenstrøm Y.H.
      • Eijsink V.G.H.
      Efficient separation of oxidized cello-oligosaccharides generated by cellulose degrading lytic polysaccharide monooxygenases.
      ,
      • Agger J.W.
      • Isaksen T.
      • Várnai A.
      • Vidal-Melgosa S.
      • Willats W.G.T.
      • Ludwig R.
      • Horn S.J.
      • Eijsink V.G.H.
      • Westereng B.
      Discovery of LPMO activity on hemicelluloses shows the importance of oxidative processes in plant cell wall degradation.
      ). Product formation in reactions with chitinous substrates was assessed by hydrophilic interaction liquid chromatography, using an UPLC (Infinity 1290; Agilent Technologies) equipped with an Acquity BEH Amide Column (130 Å, 1.7 μm, 2.1 × 150 mm) combined with a VanGuard Precolumn (Waters Corporation). The samples were adjusted to 74% acetonitrile prior to analysis, and oxidized chito-oligosaccharides were separated using a 12 min gradient as reported earlier (
      • Loose J.S.
      • Forsberg Z.
      • Kracher D.
      • Scheiblbrandner S.
      • Ludwig R.
      • Eijsink V.G.H.
      • Vaaje-Kolstad G.
      Activation of bacterial lytic polysaccharide monooxygenases with cellobiose dehydrogenase.
      ). For the identification of produced native and oxidized chito-oligosaccharides, standards were produced as previously described (
      • Loose J.S.M.
      • Forsberg Z.
      • Fraaije M.W.
      • Eijsink V.G.H.
      • Vaaje-Kolstad G.
      A rapid quantitative activity assay shows that the Vibrio cholerae colonization factor GbpA is an active lytic polysaccharide monooxygenase.
      ). To quantify conversion yields in the chitin degradation assays, soluble products (GlcNAc and GlcNAcGlcNAc1A) were analyzed using a Dionex Ultimate 3000 RSLC system (Dionex) equipped with a Rezex RFQ-Fast Acid H+ (8%) column (7.8 × 100 mm; Phenomenex) operated at 85 °C (
      • Mutahir Z.
      • Mekasha S.
      • Loose J.S.M.
      • Abbas F.
      • Vaaje-Kolstad G.
      • Eijsink V.G.H.
      • Forsberg Z.
      Characterization and synergistic action of a tetra-modular lytic polysaccharide monooxygenase from Bacillus cereus.
      ). Data were acquired and analyzed using Chromeleon, version 7.2.9 (Thermo Fisher Scientific).
      Oxidized products generated from chitin were also analyzed with MALDI-TOF MS using an Ultraflex MALDI-TOF/TOF instrument (Bruker Daltonics) equipped with a 337 nm nitrogen laser. The samples were mixed in a 1:1 ratio with a matrix solution consisting of 9 mg ml−1 2,5-dihydroxybenzoic acid and 30% acetonitrile and subsequently applied on an MTP 384TF ground steel target plate (Bruker Daltonics). Data acquisition and analysis were performed using FlexControl, version 3.4 and FlexAnalysis, version 3.4 (Bruker Daltonics), respectively.

      Data availability

      All data are contained within the article. The crystal structure of AfAA11A has been deposited in the Protein Data Bank under the accession code 7P3U.

      Conflict of interest

      The authors declare that they have no conflicts of interest with the contents of this article.

      Author contributions

      D. M. P., A. V., and V. G. H. E. conceptualization; F. G. S. and Å. K. R. formal analysis; F. G. S., Å. K. R., S. M., D. M. P., and A. V. investigation; F. G. S., Å. K. R., and V. G. H. E. writing–original draft; Å. K. R., S. M., D. M. P., A. V., and V. G. H. E. writing–review & editing; F. G. S. and Å. K. R. visualization; S. M., D. M. P., A. V., and V. G. H. E. supervision; A. V. and V. G. H. E. project administration; V. G. H. E. funding acquisition.

      Funding and additional information

      This work was supported by the following project grants from the Research Council of Norway: 243663 (to V. G. H. E., A. V., and D. M. P.), 301022 (to Å. K. R.), 297907 (to V. G. H. E., A. V., and F. G. S.), 262853 (to V. G. H. E.), and 247001 (to S. M. and V. G. H. E.).

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